INTRODUCTION.

To determine transfection efficiency, positive control standard curves were for­mulated using relative luminescence data. However, due to significant incon­sistencies among the standard curves produced transfection efficiencies were not calculated. Instead, optimal conditions were determined using the lumi­nescence signal readings of each of the complexes. Furthermore, data provided by the zetasizer was used to compare complex sizes and determine whether or not HA had stabilized the complexes. Similar size measurements were repeated for complexes in foams. In addition, DNA concentration in the releasate deter­mined by the nanodrop was used to measure cumulative release of the com­plexes from the scaffold over the span of 9 days.

RESULTS.

Optimizing Transfection Effi  ciency in vitro.

After thorough analysis of the relative luminescence data obtained from each experiment, the optimal condition for cell transfection was determined. Both fresh (data not shown) and lyophilized results showed that complexes made in DPBS with a PEI:pDNA charge ratio of 12:1 that were added to IMDF cells transfected the greatest number of cells (Figure 1 and S1). This was made evi­dent through the positive relative luminescence signal. Complexes without HA for both days 0 and day 1 had relative luminescence units (RLU’s) greater than 50 while all complexes containing HA had RLU values of 0.

However, according to the size distribution graphs, only those complexes con­taining HA remained stable over time (Figure 2). Between day 0 and day 1 of reconstituting lyophilized complexes, all complexes with HA maintained a size of roughly 200 nm, whereas, the complex condition not containing HA aggre­gated over time from 521.6 nm to 997.6 nm (Table S1 and S2). Zeta potential values also show that complexes containing HA had a net negative charge while

Figure 1. Luminescence signal of PEI:HA:pDNA complexes at day 1 (24h aft er be­ing reconstituted). Only complexes without HA in DPBS produced positive lumi­nescence readings.

Figure 2. Size distribution of lyophilized complexes at day 1. Complexes containing HA remained stable maintaining relatively small sizes while complexes without HA appeared to be greater in size.

Measuring the Release Kinetics in vitro.

Pore size, release kinetics, and cellular transfection ability were determined to further characterize the complexes and PUR foams. A scanning electron micro­scope image of the foam showed 200-500 um sized pores with an aggregated amount of complexes in the center (Figure S2). Aggregated complexes are less likely to transfect cells because of their large sizes, therefore, an even distribu­tion of complexes throughout the scaffold is more favorable.

Figure 3.Cumulative release of complexes from PUR foam over nine days. No intial burst was observed, but instead a sustained release over time.

Furthermore, Figure 3 shows that for complexes made in DPBS with a PEI:HA:pDNA charge ratio of 12:0:1, there was a sustained release over the course of roughly a week. No initial burst was observed which ensures that the complexes will be released at a constant rate over time.

To ensure that complexes being released from the foam were not only able to escape from the scaffold but also able to transfect cells following release, part of the releasate was added to IMDF cells and a luminescence reading was taken (Figure S3). The three replicates of the same PUR scaffold incubated in PBS were unable to transfect any cells as indicated by the values near 0 for RLU’s (Figure S3). This was compared with the lyophilized complexes (while still a powder, before being incorporated into PUR foams) which successfully trans­fected a greater number of cells and therefore had RLU’s surpassing 40.

Finally, size distribution of the complexes in the releasate and lyophilized com­plexes was measured (Figure S4). Although complexes in the releasate were unable to transfect cells while lyophilized complexes were, it was determined that all complexes were roughly the same size ranging from 740 nm to 990 nm (Table S3).

DISCUSSION.

Complexes made in DPBS media with a PEI:HA:pDNA charge ratio of 12:0:1, were determined to be the optimal complexes for cell transfection in vitro. These unique complexes demonstrated the most reasonable characteristics compared with other observed conditions. Complexes made in OPTI-MEM aggregated a large amount over a period of 24 hours and were therefore unable to penetrate cell membranes (data not shown). This aggregation could have been due to the interactions between the PEI in the complexes and the serum proteins present in OPTI-MEM media [3]. On the other hand, complexes made in DPBS were smaller in size primarily because of the lack of serum proteins in this media. It is important to consider this aggregation issue when complexes will be incor­porated in vivo. Here, serum protein-based blood will be one of the main body fluids with which complexes will come into contact, and aggregation between PEI and this abundance of serum proteins must be prevented to allow for com­plexes to penetrate cell membranes.

Furthermore, complexes with HA demonstrated qualities not suitable for cell transfection. As established by theAmerican Society of Gene and Cell Therapy, HA did stabilize all complexes over time by preventing drastic aggregation [3]. However, HA also made all complexes have a net negative charge which pre­vented them from being engulfed by cells. As a result all complexes containing HA were unable to transfect any cells even though their size distribution range was a lot smaller and they were more stable than complexes without HA. A smaller ratio could possibly be used to decrease the influence of HA on the net charge of complexes while still maintaining the stabilizing characteristics of HA.

Despite slight size instability and a small positive zeta potential value, com­plexes with the PEI:HA:pDNA ratio of 12:0:1 were chosen for incorporating into PUR foams. Several challenges arose during this process as well. First of all, SEM images show that complexes remained in clumps once inside the foam. Unfortunately, none of the complexes were spread evenly throughout the scaf­folds in any of the three replicates produced. Secondly, although a sustained cumulative release was observed for the complexes from the scaffold, none of the released complexes transfected any cells in vitro. This led to the hypothesis that the lyophilization process had altered the characteristics of the complexes, making them incapable of transfection. However, lyophilized complexes were tested under the same conditions of the complexes from the releasate and rela­tive luminescence readings show that the lyophilized complexes were still active and capable of transfection.

As a result, it was proposed that the size of the aggregated complexes in the scaffold was the characteristic intervening with the transfection process. Yet, looking at size distribution data, increased aggregation was not observed for any of the complexes in the releasate compared to lyophilized complexes at Day 1. Therefore, if neither freeze-drying nor size were the issue, an alternative ex­planation for the lack of transfection could be that a chemical reaction in the formation of the PUR scaffold adversely affected the complexes. Further steps could be taken to encapsulate complexes in microspheres before incorporation into the PUR scaffold to prevent degradation by any reaction. This perhaps could ensure efficient gene delivery and lead to successful skin regeneration.

ACKNOWLEDGMENTS.

I would like to thank Dr. Guelcher, Elizabeth Adolph, and The School for Science and Math at Vanderbilt for giving me the opportunity to conduct this research and for all the help, guidance, and sup­port. The project described was supported by Award Number R25RR024261 from the National Center For Research Resources. The content is solely the re­sponsibility of the authors and does not necessarily represent the official views of the National Center For Research Resources or the National Institutes of Health.

REFERENCES.

B. Li, et al., Biomaterials30: 3486-3494 (2009).

T. Ito, et al., Journal of Controlled Release 112(3): 382-388 (2006).

T. Ito, et al., Biomaterials2912-2918 (2010).

W.T. Godbey, et al., Journal of Controlled Release60: 149-160 (1999).